This research was supported by the Natural Sciences and Engineering Research Council of Canada (05239 to P.B.S.), Canadian Foundation for Innovation/Ontario Research Fund (to P.B.S.), Prostate Cancer Fight Foundation - Telus Ride for Dad (to P.B.S.) and Ontario Graduate Scholarship (to Y.J.C. and N.S.).

1. Polymerase chain reaction (PCR)-mediated site-directed mutagenesis for the incorporation of Cys into a bacterial pET-28a expression vector.
<ol>
	<li>Determine the concentration of the pET-28a vector (i.e. double stranded DNA) using a ultraviolet (UV) extinction coefficient of 0.020 (&#956;g/mL) cm-1 at 260 nm.</li>
	<li>Synthesize a pair of complementary mutagenic primers for each Cys mutation such that i) there are a minimum of 15 nucleotides complementary to the template prior to the first base mismatch and 15 nucleotides complementary to the template after the final base mismatch, ii) total primer length does not exceed 45 nucleotides, and iii) a guanine or cytosine is located at the first and last nucleotide position of each primer (Table 1). Ensure primer synthesis is performed using a 0.025 &#956;mol scale and cartridge purification.</li>
	<li>Using a high fidelity DNA polymerase, set up two 20 &#181;L PCR reaction mixtures: one containing the forward primer and the second containing the reverse primer. Prepare each mixture to contain final concentrations of 1x&#160;PCR buffer with 1.5 mM MgCl2, 0.2 mM dNTPs, 0.5 &#956;M primer, 0.4 &#956;L DMSO, 1.25 ng/&#956;L template DNA, 0.02 U/&#956;L high fidelity DNA polymerase.</li>
	<li>Thermally cycle the separate mixtures using a three-step protocol: 98 &#176;C for 30 s (denaturing), 53 - 56 &#176;C for 30 s (annealing), 72 &#176;C for 30 s kilobase(kb)-1 (extension) of the template DNA. Repeat the temperature program for 5 cycles and add a final 72 &#176;C extension step for 7.5 min.</li>
	<li>After the initial PCR with forward and reverse primers in separate tubes, combine the products into a single tube (i.e. 40 &#956;L total) and continue the PCR reaction for an additional 20 cycles using the same cycling parameters as described in step 1.4.</li>
	<li>Electrophorese 15 &#956;L of the PCR reaction mixture on a 1% (w/v) agarose gel using 0.5x Tris, acetic acid, ethylene diamine tetra acetic acid (EDTA) running buffer (TAE). As controls, electrophorese an equivalent amount of template DNA which has not been amplified by PCR and an aliquot of reference DNA ladder which contains marker bands both greater and less than the expected PCR product size.</li>
	<li>After electrophoresis at 120 V for 40 min, submerge the gel in water containing 0.5 &#956;g/mL ethidium bromide and shake for 30 min at room temperature. Confirm the full-length template has been amplified by the mutagenic primers as an increase in the relative ethidium bromide fluorescent intensity of the amplified band compared to the control template band under UV light (302 nm).
	<ol>
		<li>If no amplification is apparent, repeat the PCR after adjusting annealing temperature in 0.5 &#176;C increments between the 53 - 56 &#176;C temperature range.</li>
	</ol>
	</li>
	<li>Upon confirmation of amplification of the template by the mutagenic primers, treat the remaining ~25 &#956;L of the PCR reaction mixture with the DpnI restriction enzyme to digest the methylated template DNA. Use DpnI at 0.5 &#181;L (10 units) per 25 &#956;L PCR reaction mixture and a final concentration of 1&#215; DpnI reaction buffer. Incubate for 2.5 h at 37 &#176;C.</li>
	<li>Following template digestion, add ~5 - 10 &#956;L of the digested mixture to 100 &#181;L of heat shock competent DH5&#945; Esherichia coli cells in a 1.75 mL microcentrifuge tube. Incubate the cell-DNA mixture on ice for 60 min.</li>
	<li>Heat shock the cell-DNA mixture in the microcentrifuge tube at 42 &#176;C for 45 s on a dry heat block. After incubating the mixture on ice for 3 min, add 900 &#956;L of room temperature Luria-Bertani broth (LB) to the cells and transfer the total cell suspension into a sterile 14 mL round-bottom tube.</li>
	<li>Incubate the cell suspension at 37 &#176;C for 90 min with constant shaking at 190 rpm.</li>
	<li>Subsequently, transfer the cell suspension back into a 1.75 mL microcentrifuge tube and centrifuge at 10,000 x g for 5 min at room temperature.</li>
	<li>After the centrifugation, remove 900 &#956;L of the supernatant and resuspend the bacterial cells by gentle pipetting in the remaining 100 &#956;L of LB.</li>
	<li>Transfer the resultant concentrated cell suspension onto an LB-agar plate containing antibiotic which is selective for the expression vector (i.e. 60 &#956;g/mLKanamycin). Aseptically spread the suspension evenly on the agar plate and incubate for ~16 h at 37 &#176;C.</li>
	<li>The following day, inoculate a single colony from the plate into 5 mL of liquid LB containing the antiobiotic selection pressure (i.e. 60 &#956;g/mL Kanamycin). Grow the liquid culture overnight at 37 &#176;C with constant shaking at 37 &#176;C.</li>
	<li>Isolate and purify the propagated plasmid from the E. coli cells using a commercially available kit based on the alkaline lysis procedure 45.</li>
	<li>Confirm the mutation of interest is present and in the proper reading frame by Sanger DNA sequencing of the plasmid 46.</li>
</ol>
2. Uniform 15N-labeled protein expression in BL21 &#916;E3 Escherichia coli .
NOTE: Different recombinant proteins require different expression conditions. The following is the optimized procedure for expression of the human STIM1 EFSAM protein.
<ol>
	<li>Transform the expression vector harboring the Cys mutations (i.e. pET-28a-EFSAM) into BL21 &#916;E3 codon (+) heat shock competent cells and plate on LB-agar plates containing the antibiotic selection pressure as described in steps 1.9-1.14) with the following modifications: directly plate a 150 &#956;L aliquot out of the ~1,000 &#956;L total cell suspension on the LB-agar plate without the need for concentrating the cells in a microcentrifuge tube by centrifugation.</li>
	<li>The following day, aseptically transfer a single colony into a 200 mL Erlenmeyer flask containing 20 mL of LB supplemented with the appropriate antibiotic (i.e. 60 &#956;g/mL kanamycin for pET-28a-EFSAM). Grow this liquid starter culture overnight (i.e. ~16 h) at 37 &#176;C with constant shaking at ~ 190 rpm.</li>
	<li>On the same day as step 2.2, prepare M9 medium for 15N-labeled protein expression by autoclaving 1 L of M9 buffer salts (i.e. 42 mM Na2HPO4, 22 mM KH2PO4, 8.6 mM NaCl, pH 7.4) in a 4 L Erlenmeyer flask. Once cool, filter a mixture of 20% (w/v) D-glucose, 1 M CaCl2, 1 M thiamine, 1 M MgSO4, 1 mg/mL biotin and 0.2 g/mL15N-NH4Cl through a 0.2 &#956;m sterile syringe filter into the 1 L sterile M9 salt solution so that the final concentrations of these components are 0.2% (w/v) D-glucose, 100 &#956;M CaCl2, 50 &#956;M thiamine, 1 mM MgSO4, 1 &#956;g/mL biotin and 1 mg/mL15N-NH4Cl.</li>
	<li>The next day, aseptically transfer the 20 mL overnight liquid starter culture into a 50 mL sterile conical tube and centrifuge at 2,400 &#215;g for 15 min to pellet the cells.</li>
	<li>After decanting the LB medium, resuspend the resulting cell pellet in 10 mL of M9 minimal medium and transfer the resuspended pellet mixture into 1 L of M9 minimal medium with antibiotic (i.e. 60 &#956;g/mL kanamycin).</li>
	<li>Grow the 1 L of M9 minimal medium containing the bacterial starter culture at 37 &#176;C and ~190 rpm constant shaking until the optical density at 600 nm (OD600) reaches ~0.6-0.8.</li>
	<li>When the specfified OD600 range is reached, add 200 &#956;M of isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) to induce protein expression.</li>
	<li>After IPTG addition, continue incubating the cells for protein expression at ambient temperature with constant shaking at ~190 rpm for ~16 h (i.e. overnight).</li>
	<li>The next day, harvest the bacteria by centrifugation at ~10,000 &#215;g, 4 &#176;C for 30 min.</li>
	<li>Decant the LB and transfer the cell pellet into a 50 mL conical tube. Store the pellet at -80 &#176;C until purification.</li>
</ol>
3. Purification of recombinant protein from E. coli.
NOTE: Different recombinant proteins require distinct purification procedures. The following is the protocol for 6&#215;His-tagged EFSAM purification from inclusion bodies expressed from the pET-28a construct.
<ol>
	<li>Manually homogenize the frozen bacterial cell pellet in 6 M guanidine-HCl, 20 mM Tris-HCl (pH 8) and 5 mM &#946;-mercaptoethanol using a motorized 10 mL transfer pipette. Add approximately 40 mL of guanidine-HCl per 5 mL of wet cell pellet for this step.</li>
	<li>Following a 90 min incubation at ambient temperature with constant rotation in a hybridization oven, centrifuge the mixture at ~15,000 &#215;g, 8 &#176;C for 40 min to separate the insoluble cell debris (i.e.&#160;pellet) from the soluble protein mixture (i.e. supernatant).</li>
	<li>Add 750 &#181;L of a 50% (v/v) Ni2+-nitrilotriacetic acid agarose bead slurry to the clarified lysate and incubate for another 90 min at room temperature with inversion in a hybridization oven.</li>
	<li>Subsequently, capture the 6&#215;His-tagged protein bound to the Ni2+ by collecting the agarose beads in a gravity flow protein purification column. Allow the lysate to flow through the column completely prior to moving to step 3.5.</li>
	<li>Wash the collected beads three times with 10 mL of 6 M urea, 20 mM Tris-HCl pH 8 and 5 mM &#946;-mercaptoethanol. Ensure that the entire 10 mL passes through the column prior to each subsequent 10 mL wash.</li>
	<li>Elute the proteins in a series of 2 mL fractions using 6 M urea, 20 mM Tris-HCl pH 8, 300 mM imidazole, and 5 mM &#946;-mercaptoethanol with a 90 s incubation time between fractions. Ensure that the entire 2 mL passes through the column prior to each subsequent elution step.</li>
	<li>At this stage, confirm the protein of interest is present in the eluted fractions by Coomassie blue-stained sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) using the method of Laemmli 47. Assess the protein size, quantity and purity by comparison against standard molecular weight marker bands which are both less than and greater than the expected molecular weight of the protein of interest.</li>
	<li>Pool the eluted protein fractions into a dialysis membrane with a 3,500 Da molecular weight cutoff and incubate in 1 L refolding buffer (20 mM Tris, 300 mM NaCl, 1 mM DTT, 5 mM CaCl2, pH 8) at 4 &#176;C overnight while the buffer is being stirred by a magnetic stirrer.</li>
	<li>After ~16 h of refolding time, add ~1 U of thrombin per mg of protein directly to the dialysis bag and incubate at 4 &#176;C for an additional ~24 h.</li>
	<li>Verify the extent of the 6&#215;His tag cleavage by Coomassie-blue staining of ~15 &#956;L protein aliquots taken from the dialysis bag before and after incubation with thrombin which are electrophoresed on denaturing polyacrylamide gels (SDS-PAGE) using the method of Laemmli 47. If a ~2 kDa shift in the migration is observed corresponding to the molecular weight of the cleaved 6&#215;His tag, continue to step 3.11; if a fraction of undigested protein remains that is detectable by Coomassie-blue staining, add ~0.2 U of thrombin per mg of protein directly to the dialysis bag and incubate at 4 &#176;C for an additional ~24 h.</li>
	<li>Use size exclusion or ion exchange chromatography to further purify the protein. For anion exchange chromatography of EFSAM, remove the protein solution from the dialysis bag and concentrate ~10-fold using an ultrafiltration centrifugal concentrator with a 10,000 Da molecular weight cutoff. Subsequently, re-dilute the solution ~20-fold in a NaCl-free buffer (20 mM Tris, 5 mM CaCl2, 1 mM DTT, pH 8).</li>
	<li>Equilibrate a prepacked anion exchange column with 10 column volumes of the NaCl-free buffer described in step 3.11. Equilibrate using buffer loaded into a luer-lock syringe containing no air bubbles by pushing the solution through the column in a dropwise manner and avoiding syringe pressures which cause steady streams of solution exiting the column. Use a strong anion exchanger (e.g. crosslinked agarose with quaternary ammonium functional groups).</li>
	<li>Load the protein solution diluted into NaCl-free buffer (step 3.11) onto the column as described in step 3.12.</li>
	<li>Elute the proteins&#160;in a gradient [i.e. 0 - 60% (v/v)] of increasing NaCl buffer (20 mM Tris, 1 M NaCl, 5 mM CaCl2, 1 mM DTT, pH 8) using a two-pump fast protein liquid chromatography (FPLC) system. Set the FPLC system to collect ~1-1.5 mL fractions and monitor the protein elution profile using UV 280 nm absorbance and a flow rate 0.5 mL/min.</li>
	<li>Identify the elution peaks and fractions containing the protein of interest as well as protein purity by Coomassie blue-stained SDS-PAGE gels using the method of Laemmli 47.</li>
	<li>Pool fractions showing &#62; 95% (i.e. taken as fractions which show only a single protein band on Coomassie-blue stained gels) into a dialysis bag and exchange into experimental buffer of interest by dialysis as described in step 3.8.</li>
</ol>
4. Chemical attachment of glucose-5-MTS to protein by dialysis.
<ol>
	<li>Prepare a 55 mM stock solution of N-(&#946;-D-glucopyranosyl)-N&#39;-[2-methanethiosulfonyl)ethyl]urea (glucose-5-MTS) by dissolving 10 mg of the compound in 500 &#956;l of 100% (v/v) DMSO. Store unused glucose-5-MTS solubilized in DMSO at -20 &#176;C.</li>
	<li>Prepare the protein sample for modification by dialyzing 1.5 mL of ~60 &#956;M protein into 1 L of modification buffer made up of 20 mM MOPS, 150 mM NaCl, 5 mM CaCl2 and 0.1 mM TCEP, pH 8.3. Use a dialysis membrane molecular weight cutoff which is smaller than the size of protein being modified (e.g. use a 3,500 Da cutoff for the ~17,500 Da EFSAM).</li>
	<li>After 24 h at 4 &#176;C, transfer the sample from the dialysis bag into a microcentrifuge tube. Add the DMSO-solubilized glucose-5-MTS to a final concentration of 2 mM.</li>
	<li>Incubate the sample in the dark for 1 h at ambient temperature. During the 1 h incubation period, mix the solution by gentle tapping of the tube every 10 min.</li>
	<li>Subsequently, re-exchange the protein into the final experimental buffer containing no reducing agent by dialysis at 4 &#176;C as described in step 4.2 or by centrifugal ultrafiltration. For the ultrafiltration procedure, concentrate the ~1.5 mL protein sample to &#60; 0.5 mL and subsequently dilute in the same concentrator with the experimental buffer. Repeat this concentration-dilution step two additional times so that the total exchange is a minimum of 30&#215;30&#215;30 = 27,000-fold. For EFSAM, use 20 mM Tris, 150 mM NaCl, 5 mM CaCl2, pH 7.5 as the experimental buffer.</li>
	<li>Prepare the sample for electrospray ionization mass spectrometry by dialysis or ultrafiltration exchange as described in steps 4.2 and 4.5, respectively, into 25 mM ammonium bicarbonate or 25 mM ammonium acetate. If using dialysis, ensure you exchange at least three times to remove any residual NaCl and CaCl2 salts.</li>
	<li>Determine the accurate mass (i.e. &#177; 1 Da) of the protein of interest using electrospray ionization mass spectrometry 48,49. Expect each covalent glucose addition to a Cys thiol via the methanethiosulfonate chemical to add 281.3 Da to the protein mass (i.e. add 360.4 Da for the glucose-5-MTS and subtract 79.1 Da for the CH3SO2 leaving group during covalent attachment).</li>
</ol>
5. Solution NMR assessment of modification efficiency and structural perturbations.
<ol>
	<li>Ensure the concentration of the modified protein is &#62; 100 &#956;M after the glucose attachment and final buffer exchange. For EFSAM, estimate protein concentration using a UV extinction coefficient at 280 nm of 1.54 (mg mL-1) cm-1.</li>
	<li>Supplement the protein solution with 60 &#956;M of 4,4-dimethyl-4-silapentane-1-sulfonic acid (DSS) for shimming and pulse calibration and 10% (v/v) D2O for the signal lock. For high signal-to-noise use 600 &#956;L samples in frequency-matched 5 mm NMR tubes, inserted into a minimum 600 MHz spectrometer equipped with a triple resonance HCN cryogenic probe.</li>
	<li>Collect standard 1H-15N HSQC spectra as previously detailed 50,51 at temperature, 1H and 15N sweep widths, transient and increment settings suitable for the particular sample. For EFSAM spectra, use 20 &#176;C, 256 1H transients, 64 15N dimension increments and 1H and 15N sweep widths set to 8,000 and 1,800 Hz, respectively.</li>
	<li>Following the acquisition of the glycosylated protein spectrum, add dithiothreitol (DTT) to the NMR sample from a 1 M stock to a final concentration of 15 mM. The DTT removes the glucose moiety from the protein by reduction of the disulfide-mediated attachment.</li>
	<li>Acquire a second 1H-15N HSQC under these reduced/unmodified conditions, providing a reference spectrum to assess modification efficiency and structural perturbations caused by the glucose attachment.</li>
	<li>Process the NMR data using NMRPipe as detailed previously 52. Ensure that processing minimally includes data conversion, phasing, solvent suppression, Fourier transform and initial visualization of the spectra.</li>
	<li>Assess modification efficiency by measuring the amide peak intensities and chemical shift values in the modified and reduced spectra using the NEASY plugin on CARA 53. Ensure to evaluate the peak intensity of the Cys amide both in the glucose attached and reduced spectra. If the Cys amide cannot be reliably located in both spectra, use the intensities of residues adjacent to the Cys as a readout.</li>
	<li>Calculate the efficiency as the intensity of the amide from the Cys-modified spectrum divided by the intensity of the amide from the Cys-reduced (i.e. DTT-treated) spectrum, multiplied by 100: 
	<img alt="Equation 1" src="/files/ftp_upload/56302/56302eq1.jpg" />, where IM is the intensity of the amide in the Cys-modified spectrum and IR is the intensity of the amide in the Cys-reduced spectrum. Alternatively, evaluate the mean efficiency over several amide peaks: 
	<img alt="Equation 2" src="/files/ftp_upload/56302/56302eq2.jpg" />, where Efficiencyi is the separately determined efficiency calculated for each residue, i, and n is the total number of residues used in the calculation.</li>
	<li>Calculate chemical shift perturbations (CSP) from the chemical shift differences between the two spectra observed in the 15N and 1H dimensions of each peak and normalizing for the larger 15N chemical shift range using the following equation: 
	<img alt="Equation 3" src="/files/ftp_upload/56302/56302eq3.jpg" />, where &#916;H is the ppm change in the proton dimension and &#916;N is the ppm change in the nitrogen dimension.</li>
</ol>

Protein glycosylation is a post-translational modification where sugars are covalently attached to polypeptides primarily through linkages to amino acid side chains. As many as 50% of mammalian proteins are glycosylated 54, where the glycosylated proteins can subsequently have a diverse range of effects from altering biomolecular binding affinity, influencing protein folding, altering channel activity, targeting molecules for degradation and cellular trafficking, to name a few (reviewed in33). The important role of glycosylation in mammalian physiology is evident by the several hundreds of proteins evolved to build the full diversity of mammalian glycan structures 33. Altered N- and O-glycosylation patterns have been associated with numerous disease states including prostate (increased and decreased), breast (increased and decreased), liver (increased), ovarian (increased), pancreatic (increased) and gastric cancers (increased) 55. Furthermore, glycosylation of Tau, huntingtin, &#945;-synuclein has been found to regulate toxicity of these proteins associated with Alzheimer&#39;s, Huntington&#39;s and Parkinson&#39;s diseases 56, and a group of congenital disorders of glycosylation have been identified resulting from heritable defects in enzymes which mediate glycosylation 54. Thus, understanding the precise biophysical, biochemical and structural effects of glycosylation has the potential to tremendously impact our understanding of protein regulation and function in health and disease.
The ten sugar building blocks which lead to the diversity of glycan structures found in the mammalian glycome include fucose, galactose, glucose, N-acetylgalactosamine, N-acetylglucosamine, glucuronic acid, iduronic acid, mannose, sialic acid and xylose 33. While N-glycosylation invariably links an N-acetylglucosamine sugar directly to the protein, O-glycosylation can result from any of N-acetylgalacotose, N-acetylglucosamine, xylose, fucose, glucose or galactose covalently linked to the polypeptide. To begin to understand how these sugars immediately adjacent to the protein surface affect the biophysical and structural properties, we describe herein an approach to site-selectively attach sugars to Cys residues via the thiols engineered into the protein sequence. Here, the residues that are endogenously glycosylated are replaced by Cys and modified in vitro via a simple chemical approach. In this manner, single and multiple glycosylation sites may be assessed to tease out the contribution of each specific site as well as the cumulative modifications to the folding and stability as well as the overall structure and function of the protein.
Recently, this approach was successfully used with EFSAM to individually and cumulatively assess the role of the Asn131 and Asn171 N-glycosylation sites 42. Mutation to Cys and covalent attachment of glucose to the Asn131 or Asn171 sites revealed a decreased Ca2+ binding affinity and suppressed stability. When the two sites were simultaneously modified with the glucose attachment, the decreases in binding affinity and stability were potentiated leading to enhanced oligomerization propensity in vitro. Structurally, the approach described herein showed that the Asn131 or Asn171 modifications mutually perturb the core &#945;8 helix located on the SAM domain, immediately adjacent to the EF-hand pair. This structural analysis expounds how glucose modifications on the surface of the protein lead to a converging and potentiated structural change within the EF-hand:SAM interface which ultimately destabilizes the protein and enhances SOCE 42.
While the application of this site-selective approach helped shed light on how a monosaccharide close to the surface of EFSAM effects folding, stability and structure, this procedure can easily be modified to attach longer carbohydrates specific to ER, Golgi and PM localization (i.e. glycosylation states of different maturity), provided there is a reliable source for these carbohydrates containing functional groups which can link to thiols such as MTS. MTS is preferable since the thiol modification is reversible using a reducing agent and a reference spectrum can be easily acquired. This approach can also be adapted to link other post-translational moieties to the protein such as lipids. At the same time, there are several limitations to this approach which should be considered. First, the method relies on mutation of glycosylation sites to Cys which may affect structure, stability and folding even in the absence of any glucose modification. Similarly, native Cys residues in the protein must also be mutated to prevent glucose attachment at non-glycosylated sites. Additionally, the addition of Cys residues often promotes inclusion body formation in bacteria due to Cys crosslinking and misfolding, making purification more challenging. Nevertheless, this site-selective Cys-crosslinking approach described herein provides a controlled means to tease out the structural, biochemical and biophysical effects of specific glycosylation sites in experiments which require high levels of homogeneous protein. The effects of non-native Cys residues on the structure, stability and folding can be simply ascertained in the absence of any modifications by comparison to wild-type protein attributes 42. Taken together with functional data obtained in eukaryotic cells which express modification-blocking mutant versions of the protein (e.g. Asn-to-Ala), the presently described approach will yield new insights into the structural mechanisms of protein regulation by post-translational modifications.

Store operated calcium (Ca2+) entry (SOCE) is the major pathway by which immune cells take up Ca2+ from the extracellular space into the cytosol. In T lymphocytes, T cell receptors located on the plasma membrane (PM) bind antigens which activate protein tyrosine kinases (reviewed in 1,2,3). A phosphorylation cascade leads to the activation of phospholipase-&#947; (PLC&#947;) which subsequently mediates the hydrolysis of membrane phosphatidylinositol 4,5-bisphosphate (PIP2) into diacylglycerol and inositol 1,4,5-trisphosphate (IP3). IP3 is a small diffusible messenger which binds to IP3 receptors (IP3R) on the endoplasmic reticulum (ER) thereby opening this receptor channel and permitting Ca2+ to flow down the concentration gradient from the ER lumen to the cytosol (reviewed in 4). Receptor signaling from G protein coupled and tyrosine kinase receptors in a variety of other excitable and non-excitable cell types lead to the same production of IP3 and activation of IP3Rs.
Due to the finite Ca2+ storage capacity of the ER, the IP3-mediated release and resultant increase in cytosolic Ca2+ is only transient; however, this depletion of the ER luminal Ca2+ profoundly effects stromal interaction molecule-1 (STIM1), a type-I transmembrane (TM) protein mostly found on the ER membrane 5,6,7. STIM1 contains a lumen-oriented Ca2+ sensing domain made up of an EF-hand pair and sterile &#945;-motif (EFSAM). Three cytosolic-oriented coiled-coil domains are separated from EFSAM by the single TM domain (reviewed in 8). Upon ER luminal Ca2+ depletion, EFSAM undergoes a destabilization-coupled oligomerization 7,9 which causes structural rearrangements of the TM and coiled-coil domains 10. These structural changes culminate in a trapping of STIM1 at ER-PM junctions 11,12,13,14 through interactions with PM phosphoinositides 15,16 and Orai1 subunits 17,18. Orai1 proteins are the PM subunits which assemble to form Ca2+ channels 19,20,21,22. The STIM1-Orai1 interactions at ER-PM junctions facilitate an open Ca2+ release activated Ca2+ (CRAC) channel conformation which enables the movement of Ca2+ into the cytosol from the high concentrations of the extracellular space. In immune cells, the sustained cytosolic Ca2+ elevations via CRAC channels induce the Ca2+-calmodulin/calcineurin dependent dephosphorylation of the nuclear factor of activated T-cells which subsequently enters the nucleus and begins transcriptional regulation of genes promoting T-cell activation 1,3. The process of CRAC channel activation by STIM1 23,24 via agonist-induced ER luminal Ca2+ depletion and the resulting sustained cytosolic Ca2+ elevation is collectively termed SOCE 25. The vital role of SOCE in T-cells is evident by studies demonstrating that heritable mutations in both STIM1 and Orai1 can cause severe combined immunodeficiency syndromes 3,19,26,27. EFSAM initiates SOCE after sensing ER-luminal Ca2+ depletion via the loss of Ca2+ coordination at the canonical EF-hand, ultimately leading to the destabilization-coupled self-association 7,28,29.
Glycosylation is the covalent attachment and processing of oligosaccharide structures, also known as glycans, through various biosynthetic steps in the ER and Golgi (reviewed in 30,32,33). There are two predominant types of glycosylation in eukaryotes: N-linked and O-linked, depending on the specific amino acid and atom bridging the linkage. In N-glycosylation, glycans are attached to the side chain amide of Asn, and in most cases, the initiation step occurs in the ER as the polypeptide chain moves into the lumen 34. The first step of N-glycosylation is the transfer of a fourteen-sugar core structure made up of glucose (Glc), mannose (Man), and N-acetylglucosamine (GlcNAc) (i.e. Glc3Man9GlcNAc2) from an ER membrane lipid by an oligosaccharyltransferase 35,36. Further steps, such as cleavage or transfer of glucose residues, are catalyzed in the ER by specific glycosidases and glycosyltransferases. Some proteins that leave the ER and move into the Golgi can be further processed 37. O-glycosylation refers to the addition of glycans, usually to the side chain hydroxyl group of Ser or Thr residues, and this modification occurs entirely in the Golgi complex 33,34. There are several O-glycan structures which can be made up of N-acetylglucosamine, fucose, galactose, and sialic acid with each monosaccharide added sequentially 33.
While no specific sequence has been identified as prerequisite for many types of O-glycosylation, a common consensus sequence has been associated with the N-linked modification: Asn-X-Ser/Thr/Cys, where X can be any amino acid except Pro 33. STIM1 EFSAM contains two of these consensus N-glycosylation sites: Asn131-Trp132-Thr133 and Asn171-Thr172-Thr173. Indeed, previous studies have shown that EFSAM can be N-glycosylated in mammalian cells at Asn131 and Asn171 38,39,40,41. However, previous studies of the consequences of N-glycosylation on SOCE have been incongruent, suggesting suppressed, potentiated or no effect by this post-translational modification on SOCE activation 38,39,40,41. Thus, research on the underlying biophysical, biochemical, and structural consequences of EFSAM N-glycosylation is vital to comprehending the regulatory effects of this modification. Due to the requirement for high levels of homogeneous proteins in these in vitro experiments, a site-selective approach to covalently attach glucose moieties to EFSAM was applied. Interestingly, Asn131 and Asn171 glycosylation caused structural changes that converge within the EFSAM core and enhance the biophysical properties which promote STIM1-mediated SOCE 42.
The chemical attachment of glycosyl groups to Cys thiols has been well-established by a seminal work which first demonstrated the utility of this enzyme-free approach to understanding the site-specific effects of glycosylation on protein function 43,44. More recently and with respect to STIM1, the Asn131 and Asn171 residues were mutated to Cys and glucose-5-(methanethiosulfonate) [glucose-5-(MTS)] was used to covalently link glucose to the free thiols 42. Here, we describe this approach which not only uses mutagenesis to incorporate site specific Cys residues for modification, but also applies solution nuclear magnetic resonance (NMR) spectroscopy to rapidly assess both modification efficiency and structural perturbations as a result of the glycosylation. Notably, this general methodology is easily adaptable to study the effects of either O- or N-glycosylation of any recombinantly produced protein.

<table><tbody><tr><td>Phusion DNA Polymerase</td><td>Thermo Fisher Scientific</td><td>F530S</td><td>Use in step 1.3.</td></tr><tr><td>Generuler 1kb DNA Ladder</td><td>Thermo Fisher Scientific</td><td>FERSM1163</td><td>Use in step 1.6.</td></tr><tr><td>DpnI Restriction Enzyme</td><td>New England Biolabs, Inc.</td><td>R0176</td><td>Use in step 1.8.</td></tr><tr><td>Presto Mini Plasmid Kit</td><td>GeneAid, Inc.</td><td>PDH300</td><td>Use in step 1.16.</td></tr><tr><td>BL21 DE3 codon (+) E. coli</td><td>Agilent Technologies, Inc.</td><td>230280</td><td>Use in step 2.1.</td></tr><tr><td>DH5a E. coli</td><td>Invitrogen, Inc.</td><td>18265017</td><td>Use in step 1.9.</td></tr><tr><td>0.22 mm Syringe Filter</td><td>Millipore, Inc.</td><td>SLGV033RS</td><td>Use in step 2.3.</td></tr><tr><td>HisPur Ni&lt;sup&gt;2+&lt;/sup&gt;-NTA Agarose Resin</td><td>Thermo Fisher Scientific</td><td>88221</td><td>Use in step 3.3.</td></tr><tr><td>3,500 Da MWCO Dialysis Tubing</td><td>BioDesign, Inc.</td><td>D306</td><td>Use in step 3.8, 3.16, 4.2, 4.5 and 4.6.</td></tr><tr><td>Bovine Thrombin</td><td>BioPharm Laboratories, Inc.</td><td>SKU91-055</td><td>Use in step 3.9.</td></tr><tr><td>5 mL HiTrap Q FF Anion Exchange Column</td><td>GE Healthcare, Inc.</td><td>17-5156-01</td><td>Use in step 3.11.</td></tr><tr><td>Glucose-5-MTS</td><td>Toronto Research Chemicals, Inc.</td><td>G441000</td><td>Use in step 4.1.</td></tr><tr><td>Vivaspin 20 Ultrafiltration Centrifugal Concentrators</td><td>Sartorius, Inc.</td><td>VS2001</td><td>Use in step 3.11, 4.2, 4.5 and 4.6.</td></tr><tr><td>PageRuler Unstained Broad Protein Ladder</td><td>Thermo Fisher Scientific</td><td>26630</td><td>Use in step 3.7, 3.10 and 3.15</td></tr><tr><td>HiTrap Q FF Anion Exchange Column</td><td>GE Healthcare, Inc.</td><td>17-5053-01</td><td>Use in step 3.12.</td></tr><tr><td>AKTA Pure Fast Protein Liquid Chromatrography System</td><td>GE Healthcare, Inc.</td><td>29018224</td><td>Use in step 3.14.</td></tr><tr><td>600 MHz Varian Inova NMR Spectrometer</td><td>Agilent Technologies, Inc.</td><td></td><td>Use in step 5.2 and 5.5.</td></tr></tbody></table>

targeting cysteine thiols for in vitro site-specific glycosylation of recombinant proteins

Stromal interaction molecule-1 (STIM1) is a type-I transmembrane protein located on the endoplasmic reticulum (ER) and plasma membranes (PM). ER-resident STIM1 regulates the activity of PM Orai1 channels in a process known as store operated calcium (Ca2+) entry which is the principal Ca2+ signaling process that drives the immune response. STIM1 undergoes post-translational N-glycosylation at two luminal Asn sites within the Ca2+ sensing domain of the molecule. However, the biochemical, biophysical, and structure biological effects of N-glycosylated STIM1 were poorly understood until recently due to an inability to readily obtain high levels of homogeneous N-glycosylated protein. Here, we describe the implementation of an in vitro chemical approach which attaches glucose moieties to specific protein sites applicable to understanding the underlying effects of N-glycosylation on protein structure and mechanism. Using solution nuclear magnetic resonance spectroscopy we assess both efficiency of the modification as well as the structural consequences of the glucose attachment with a single sample. This approach can readily be adapted to study the myriad glycosylated proteins found in nature.

The first step of this approach requires the mutagenesis of the candidate glycosylation residues to Cys residues which can be modifiable using the glucose-5-MTS. EFSAM has no endogenous Cys residues, so no special considerations need to be made prior to the mutagenesis. However, native Cys residues must be mutated to non-modifiable residues prior to performing the described chemistry. To minimally effect the native structure, we suggest performing a global sequence alignment of the protein of interest and determining which other residues are found most frequently at the endogenous Cys position(s). Cys mutation to these other residues which occur naturally in other organisms may have the least impact on protein structure. If the endogenous Cys residue is strictly conserved, we suggest mutating to Ser which is the most structurally similar to Cys. Figure 1 shows a typical PCR mutagenesis gel evaluating the success of the PCR reaction, with the amplified DNA sample demonstrating several-fold higher intensity than a control amount of unamplified template pET-28a DNA which was used for PCR. The next steps include template DNA digestion and transformation into E. coli for plasmid repair. After plasmid propagation in liquid culture, plasmid isolation and confirmation of the mutation(s) by sequencing, the mutated vector may be used for protein expression. Figure 2A shows a typical elution profile of EFSAM from the anion exchange column relative to increasing NaCl concentrations. Figure 2B shows the purity of EFSAM on Coomassie blue-stained SDS-PAGE gels.
After acquiring pure protein, a series of dialysis steps is used to attach the glucose moiety via the MTS reaction with the free thiol. Figure 3A shows an image of the typical setup of a small protein volume sealed in the dialysis membrane by membrane clips and contained in a large 1 L beaker containing the buffer of interest. An initial check of the success of the modification may be performed by mass spectrometry. Figure 3B shows a representative electrospray mass spectrum of EFSAM modified at a single Cys thiol. Following establishment of the protocol for a specific protein, modification efficiency and structural perturbations can be assessed from a single uniformly 15N-labeled sample. The 1H-15N-HSQC spectrum is acquired before and after the addition of the reducing agent DTT (Figure 4A). Calculations of the modification efficiency can be made via a comparison of the amide peak intensities in the modified and reduced spectra as detailed in protocol step 5.8&#160;(Figure 4B). Finally, when chemical shift assignments are known for a protein, the CSPs which correlate with the structural changes can be calculated as detailed in step 5.9&#160;(Figure 4C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56302/56302fig1.jpg" /> 
Figure 1: DNA agarose gel showing amplification check of template vector with mutagenic primers. 
The image shows a 1.0% (w/v) agarose gel with DNA marker (M), vector control (VC) and PCR-amplified template (PCR). The DNA was separated by electrophoresis at 120 V for 45 min in 0.5x&#160;TAE buffer. A total of 0.5 ng of VC was loaded, equivalent to the amount of template loaded into the PCR lane. The gel was stained using ethidium bromide (~0.5 &#956;g/mL) for 20 min prior to visualization under UV light (302 nm). The gel shows a high level of amplified DNA close to the expected size of the vector (black arrowhead). The second band in the PCR lane running between the 1,000 and 1,500 bp marker bands likely represents a non-specifically amplified PCR product. The intensity level of amplified DNA must be higher than the VC intensity level to be deemed successful. Several other DNA dyes can be used as less mutagenic, safer alternatives to ethidium bromide staining (see for example 57). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56302/56302fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56302/56302fig2.jpg" /> 
Figure 2: Typical chromatographic purification and purity check for STIM1 EF-SAM. 
(A) Anion exchange chromatography elution profile of STIM1 EF-SAM. After manual binding EF-SAM to the anion exchange column (Q FF) at basic pH and low NaCl concentration with a syringe, and AKTA FPLC (GE Healthcare) is used to elute the protein with a NaCl gradient. The elution is monitored by the AKTA using the UV 280 nm signal over a 0-60% (v/v) gradient of 1 M NaCl. (B) Coomassie blue-stained SDS-PAGE gel of elution fractions from (A). The denaturing protein gel reveals that EF-SAM elutes in two major peaks at ~250 mM and ~450 mM NaCl. The purification protocol yields &#62; 95% pure EF-SAM as evidenced by the lack of any contaminant band showing up in the Coomassie blue-stained gel. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56302/56302fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56302/56302fig3.jpg" /> 
Figure 3: Dialysis setup and confirmation of in vitro protein glycosylation. 
(A) Typical dialysis setup used for in vitro attachment of glucose to the Cys thiol via the MTS reactivity. The image shows ~1.5 mL of protein contained within dialysis tubing buffered against ~ 1 L of experimental buffer. It is important that the buffer is constantly stirred to ensure complete exchange. The image shows a microcentrifuge tube clipped to the excess dialysis tubing to prevent sinking of the dialysis bag and damage by the rotating stir bar. (B) Electrospray ionization mass spectrum of the modified Asn171Cys EF-SAM protein. Mass spectrometry is a convenient and accurate approach to assess whether the modification procedure was successful. A typical mass chromatogram is shown with the theoretical and measured masses of unmodified and modified Asn171Cys EF-SAM indicated. The majority of the sample mass corresponds to a macromolecule which is within ~1.3 Da of the expected theoretical mass of glucose-conjugated Asn171Cys EF-SAM. The data in (B) is replotted and modified from 42. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56302/56302fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56302/56302fig4.jpg" /> 
Figure 4: Solution NMR assessment of modification efficiency and structural perturbations from a single NMR sample. 
(A)1H-15N-HSQC spectral overlay of glucose-conjugated Asn131Cys EFSAM before (red crosspeaks) and after (black crosspeaks) the addition of 15 mM DTT. The overlay clearly shows several residue-specific amide chemical shift changes indicative of both modification of the protein and structural perturbations. The red box shows the location of the Asn131Cys amide. (B) Zoomed view of the 1H-15N-HSQC region containing the Asn131Cys amide. The intensity of the Asn131Cys amide peak in the modified spectrum (IM) is divided by the intensity in the reduced (unmodified) spectrum (IR) for the calculation of efficiency (shown). A calculation of the mean efficiency of several effected residues provides a better estimate of efficiency, including an error estimate. The mean efficiency is shown for Asn131Cys EFSAM based on 5 residues (i.e. 129-133). (C) Normalized chemical shift perturbations caused by glucose conjugation to the Asn131Cys EFSAM protein. The set of HSQC experiments collected on a single sample before and after supplementing with reducing agent not only provides a convenient estimate of modification efficiency by peak intensity analysis [shown in (B)], but also provides data for evaluation of the structural changes associated with the modification. Glucose conjugation causes the largest perturbations localized near position 131; however, this analysis reveals perturbations which are unexpected solely based on sequence proximity, indicating the value in this analysis. The data in (C) are replotted and modified from 42. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56302/56302fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>STIM1 mutationa</td>
			<td>directionb</td>
			<td colspan="2">DNA sequencec</td>
		</tr>
		<tr>
			<td>Asn131Cys</td>
			<td>forward</td>
			<td colspan="2">5&#8217;-GTCATCAGAAGTATACTGTTGGACCGTGGATGAGG-3&#8217;</td>
		</tr>
		<tr>
			<td>Asn131Cys</td>
			<td>reverse</td>
			<td colspan="2">5&#8217;-CCTCATCCACGGTCCAACAGTATACTTCTGATGAC-3&#8217;</td>
		</tr>
		<tr>
			<td>Asn171Cys</td>
			<td>forward</td>
			<td colspan="2">5&#8217;-CCAAGGCTGGCTGTCACCTGCACCACCATGACAGGG-3&#8217;</td>
		</tr>
		<tr>
			<td>Asn171Cys</td>
			<td>reverse</td>
			<td colspan="2">5&#8217;-CCCTGTCATGGTGGTGCAGGTGACAGCCAGCCTTGG-3&#8217;</td>
		</tr>
		<tr>
			<td colspan="4">aSTIM1 amino acid numbering based on NCBI accession AFZ76986.1.</td>
		</tr>
		<tr>
			<td colspan="4">bThe &#8216;reverse&#8217; primer corresponds to the reverse complement sequence of the &#8216;forward&#8217; primer.</td>
		</tr>
		<tr>
			<td colspan="4">cThe underlined codon triplet corresponds to the Cys mutation.</td>
		</tr>
	</tbody>
</table>
Table 1. Example oligonucleotide (primer) sequences for Asn to Cys mutagenesis within the pET-28a STIM1 EFSAM construct.

Watch this Scientific Journal Video about Targeting Cysteine Thiols for in Vitro Site-specific Glycosylation of Recombinant Proteins at JoVE.com

Targeting Cysteine Thiols for in Vitro Site-specific Glycosylation of Recombinant Proteins

Biochemical and structural analyses of glycosylated proteins require relatively large amounts of homogeneous samples. Here, we present an efficient chemical method for site-specific glycosylation of recombinant proteins purified from bacteria by targeting reactive Cys thiols.

Targeting Cysteine Thiols for in Vitro Site-specific Glycosylation of Recombinant Proteins

Stromal interaction molecule-1 (STIM1) is a type-I transmembrane protein located on the endoplasmic reticulum (ER) and plasma membranes (PM). ER-resident ...

targeting-cysteine-thiols-for-vitro-site-specific-glycosylation

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

6.5K Views.  University of Western Ontario. Biochemische en structurele analyses van geglycosyleerde eiwitten vereist relatief grote hoeveelheden van homogene monsters. Hier presenteren we een efficiënte chemische methode voor site-specifieke glycosylatie van recombinante eiwitten gezuiverd van bacteriën door zich te richten van reactieve Cys thiolen.

Video: Targeting Cysteine Thiols for in Vitro Site-specific Glycosylation of Recombinant Proteins

Synthesis of Protein Bioconjugates via Cysteine-maleimide Chemistry

The chemical linking or bioconjugation of proteins to fluorescent dyes, drugs, polymers and other proteins has a broad range of applications, such as the development of antibody drug conjugates (ADCs) and nanomedicine, fluorescent microscopy and systems chemistry. For many of these applications, specificity of the bioconjugation method used is of prime concern. The Michael addition of maleimides with cysteine(s) on the target proteins is highly selective and proceeds rapidly under mild conditions, making it one of the most popular methods for protein bioconjugation.
We demonstrate here the modification of the only surface-accessible cysteine residue on yeast cytochrome c with a ruthenium(II) bisterpyridine maleimide. The protein bioconjugation is verified by gel electrophoresis and purified by aqueous-based fast protein liquid chromatography in 27% yield of isolated protein material. Structural characterization with MALDI-TOF MS and UV-Vis is then used to verify that the bioconjugation is successful. The protocol shown here is easily applicable to other cysteine - maleimide coupling of proteins to other proteins, dyes, drugs or polymers.

Synthesis of Protein Bioconjugates via Cysteine-maleimide Chemistry

This protocol details the important steps required for the bioconjugation of a cysteine containing protein to a maleimide, including reagent purification, reaction conditions, bioconjugate purification and bioconjugate characterization.

The chemical linking or bioconjugation of proteins to fluorescent dyes, drugs, polymers and other proteins has a broad range of applications, such as the ...

Chemistry

Characterization of Glycoproteins with the Immunoglobulin Fold by X-Ray Crystallography and Biophysical Techniques

Glycoproteins on the surface of cells play critical roles in cellular function, including signalling, adhesion and transport. On leukocytes, several of these glycoproteins possess immunoglobulin (Ig) folds and are central to immune recognition and regulation. Here, we present a platform for the design, expression and biophysical characterization of the extracellular domain of human B cell receptor CD22. We propose that these approaches are broadly applicable to the characterization of mammalian glycoprotein ectodomains containing Ig domains. Two suspension human embryonic kidney (HEK) cell lines, HEK293F and HEK293S, are used to express glycoproteins harbouring complex and high-mannose glycans, respectively. These recombinant glycoproteins with different glycoforms allow investigating the effect of glycan size and composition on ligand binding. We discuss protocols for studying the kinetics and thermodynamics of glycoprotein binding to biologically relevant ligands and therapeutic antibody candidates. Recombinant glycoproteins produced in HEK293S cells are amenable to crystallization due to glycan homogeneity, reduced flexibility and susceptibility to endoglycosidase H treatment. We present methods for soaking glycoprotein crystals with heavy atoms and small molecules for phase determination and analysis of ligand binding, respectively. The experimental protocols discussed here hold promise for the characterization of mammalian glycoproteins to give insight into their function and investigate the mechanism of action of therapeutics.

We present approaches for the biophysical and structural characterization of glycoproteins with the immunoglobulin fold by biolayer interferometry, isothermal titration calorimetry, and X-ray crystallography.

Glycoproteins on the surface of cells play critical roles in cellular function, including signalling, adhesion and transport. On leukocytes, several of ...

Biochemistry

Synthesis and Bioconjugation of Thiol-Reactive Reagents for the Creation of Site-Selectively Modified Immunoconjugates

Maleimide-bearing bifunctional probes have been employed for decades for the site-selective modification of thiols in biomolecules, especially antibodies. Yet maleimide-based conjugates display limited stability in vivo because the succinimidyl thioether linkage can undergo a retro-Michael reaction. This, of course, can lead to the release of the radioactive payload or its exchange with thiol-bearing biomolecules in circulation. Both of these processes can produce elevated activity concentrations in healthy organs as well as decreased activity concentrations in target tissues, resulting in reduced imaging contrast and lower therapeutic ratios. In 2018, we reported the creation of a modular, stable, and easily accessible phenyloxadiazolyl methyl sulfone reagent &#8212; dubbed &#8216;PODS&#8217; &#8212; as a platform for thiol-based bioconjugations. We have clearly demonstrated that PODS-based site-selective bioconjugations reproducibly and robustly create homogenous, well-defined, highly immunoreactive, and highly stable radioimmunoconjugates. Furthermore, preclinical experiments in murine models of colorectal cancer have shown that these site-selectively labeled radioimmunoconjugates exhibit far superior in vivo performance compared to radiolabeled antibodies synthesized via maleimide-based conjugations. In this protocol, we will describe the four-step synthesis of PODS, the creation of a bifunctional PODS-bearing variant of the ubiquitous chelator DOTA (PODS-DOTA), and the conjugation of PODS-DOTA to the HER2-targeting antibody trastuzumab.

In this protocol, we will describe the synthesis of PODS, a phenyoxadiazolyl methyl sulfone-based reagent for the site-selective attachment of cargos to the thiols of biomolecules, particularly antibodies. In addition, we will describe the synthesis and characterization of a PODS-bearing bifunctional chelator and its conjugation to a model antibody.

Maleimide-bearing bifunctional probes have been employed for decades for the site-selective modification of thiols in biomolecules, especially antibodies. ...

Preparation of Site-Specific Cytotoxic Protein Conjugates via Maleimide-thiol Chemistry and Sortase A-Mediated Ligation

Cancer is currently the second most common cause of death worldwide. The hallmark of cancer cells is the presence of specific marker proteins such as growth factor receptors on their surface. This feature enables development of highly selective therapeutics, the protein bioconjugates, composed of targeting proteins (antibodies or receptor ligands) connected to highly cytotoxic drugs by a specific linker. Due to very high affinity and selectivity of targeting proteins the bioconjugates recognize marker proteins on the cancer cells surface and utilize receptor-mediated endocytosis to reach the cell interior. Intracellular vesicular transport system ultimately delivers the bioconjugates to the lysosomes, where proteolysis separates free cytotoxic drugs from the proteinaceous core of the bioconjugates, triggering drug-dependent cancer cell death. Currently, there are several protein bioconjugates approved for cancer treatment and large number is under development or clinical trials.
One of the main challenges in the generation of the bioconjugates is a site-specific attachment of the cytotoxic drug to the targeting protein. Recent years have brought a tremendous progress in the development of chemical and enzymatic strategies for protein modification with cytotoxic drugs. Here we present the detailed protocols for the site-specific incorporation of cytotoxic warheads into targeting proteins using a chemical method employing maleimide-thiol chemistry and an enzymatic approach that relies on sortase A-mediated ligation. We use engineered variant of fibroblast growth factor 2 and fragment crystallizable region of human immunoglobulin G as an exemplary targeting proteins and monomethyl auristatin E and methotrexate as model cytotoxic drugs. All the described strategies allow for highly efficient generation of biologically active cytotoxic conjugates of defined molecular architecture with potential for selective treatment of diverse cancers.

Here we provide detailed protocols for a site-specific labeling of proteins with cytotoxic drugs using maleimide-thiol reaction and sortase A-mediated ligation.

Cancer is currently the second most common cause of death worldwide. The hallmark of cancer cells is the presence of specific marker proteins such as ...

Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed procedure of a quantitative redox proteomics method that utilizes resin-assisted capture (RAC) in combination with tandem mass tag (TMT) isobaric labeling and liquid chromatography-tandem mass spectrometry (LC-MS/MS) to allow multiplexed stochiometric quantification of oxidized protein thiols at the proteome level. The site-specific quantitative information on oxidized cysteine residues provides additional insight into the functional impacts of such modifications. 
The workflow is adaptable across many sample types, including cultured cells (e.g., mammalian, prokaryotic) and whole tissues (e.g., heart, lung, muscle), which are initially lysed/homogenized and with free thiols being alkylated to prevent artificial oxidation. The oxidized protein thiols are then reduced and captured by a thiol-affinity resin, which streamlines and simplifies the workflow steps by allowing the proceeding digestion, labeling, and washing procedures to be performed without additional transfer of proteins/peptides. Finally, the labeled peptides are eluted and analyzed by LC-MS/MS to reveal comprehensive stoichiometric changes related to thiol oxidation across the entire proteome. This method greatly improves the understanding of the role of redox-dependent regulation under physiological and pathophysiological states related to protein thiol oxidation.

Protein thiol oxidation has significant implications under normal physiological and pathophysiological conditions. We describe the details of a quantitative redox proteomics method, which utilizes resin-assisted capture, isobaric labeling, and mass spectrometry, enabling site-specific identification and quantification of reversibly oxidized cysteine residues of proteins.

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Biology

Homogeneous Glycoconjugate Produced by Combined Unnatural Amino Acid Incorporation and Click-Chemistry for Vaccine Purposes

Genetic code expansion is a powerful tool to introduce unnatural amino acids (UAAs) into proteins to modify their characteristics, to study or create new protein functions or to have access to protein conjugates. Stop codon suppression, in particular amber codon suppression, has emerged as the most popular method to genetically introduce UAAs at defined positions. This methodology is herein applied to the preparation of a carrier protein containing an UAA harboring a bioorthogonal functional group. This reactive handle can next be used to specifically and efficiently graft a synthetic oligosaccharide hapten to provide a homogeneous glycoconjugate vaccine. The protocol is limited to the synthesis of glycoconjugates in a 1:1 carbohydrate hapten/carrier protein ratio but amenable to numerous pairs of biorthogonal functional groups. Glycococonjugate vaccine homogeneity is an important criterion to ensure complete physico-chemical characterization, thereby, satisfying more and more demanding drug regulatory agency recommendations, a criterion which is unmet by classical conjugation strategies. Moreover, this protocol makes it possible to finely tune the structure of the actual conjugate vaccine, giving rise to tools to address structure-immunogenicity relationships.

Genetic code expansion is applied for the introduction of an unnatural amino acid bearing a biorthogonal functional group on a carrier protein at a defined site. The biorthogonal function is further used for the site-selective coupling of a carbohydrate antigen to provide a homogeneous glycoconjugate vaccine.

Genetic code expansion is a powerful tool to introduce unnatural amino acids (UAAs) into proteins to modify their characteristics, to study or create new ...

Bioengineering

Chemically-blocked Antibody Microarray for Multiplexed High-throughput Profiling of Specific Protein Glycosylation in Complex Samples

In this study, we describe an effective protocol for use in a multiplexed high-throughput antibody microarray with glycan binding protein detection that allows for the glycosylation profiling of specific proteins. Glycosylation of proteins is the most prevalent post-translational modification found on proteins, and leads diversified modifications of the physical, chemical, and biological properties of proteins. Because the glycosylation machinery is particularly susceptible to disease progression and malignant transformation, aberrant glycosylation has been recognized as early detection biomarkers for cancer and other diseases. However, current methods to study protein glycosylation typically are too complicated or expensive for use in most normal laboratory or clinical settings and a more practical method to study protein glycosylation is needed. The new protocol described in this study makes use of a chemically blocked antibody microarray with glycan-binding protein (GBP) detection and significantly reduces the time, cost, and lab equipment requirements needed to study protein glycosylation. In this method, multiple immobilized glycoprotein-specific antibodies are printed directly onto the microarray slides and the N-glycans on the antibodies are blocked. The blocked, immobilized glycoprotein-specific antibodies are able to capture and isolate glycoproteins from a complex sample that is applied directly onto the microarray slides. Glycan detection then can be performed by the application of biotinylated lectins and other GBPs to the microarray slide, while binding levels can be determined using Dylight 549-Streptavidin. Through the use of an antibody panel and probing with multiple biotinylated lectins, this method allows for an effective glycosylation profile of the different proteins found in a given human or animal sample to be developed.
Introduction
Glycosylation of protein, which is the most ubiquitous post-translational modification on proteins, modifies the physical, chemical, and biological properties of a protein, and plays a fundamental role in various biological processes1-6. Because the glycosylation machinery is particularly susceptible to disease progression and malignant transformation, aberrant glycosylation has been recognized as early detection biomarkers for cancer and other diseases 7-12. In fact, most current cancer biomarkers, such as the L3 fraction of &alpha;-1 fetoprotein (AFP) for hepatocellular carcinoma 13-15, and CA199 for pancreatic cancer 16, 17 are all aberrant glycan moieties on glycoproteins. However, methods to study protein glycosylation have been complicated, and not suitable for routine laboratory and clinical settings. Chen et al. has recently invented a chemically blocked antibody microarray with a glycan-binding protein (GBP) detection method for high-throughput and multiplexed profile glycosylation of native glycoproteins in a complex sample 18. In this affinity based microarray method, multiple immobilized glycoprotein-specific antibodies capture and isolate glycoproteins from the complex mixture directly on the microarray slide, and the glycans on each individual captured protein are measured by GBPs. Because all normal antibodies contain N-glycans which could be recognized by most GBPs, the critical step of this method is to chemically block the glycans on the antibodies from binding to GBP. In the procedure, the cis-diol groups of the glycans on the antibodies were first oxidized to aldehyde groups by using NaIO4 in sodium acetate buffer avoiding light. The aldehyde groups were then conjugated to the hydrazide group of a cross-linker, 4-(4-N-MaleimidoPhenyl)butyric acid Hydrazide HCl (MPBH), followed by the conjugation of a dipeptide, Cys-Gly, to the maleimide group of the MPBH. Thus, the cis-diol groups on glycans of antibodies were converted into bulky none hydroxyl groups, which hindered the lectins and other GBPs bindings to the capture antibodies. This blocking procedure makes the GBPs and lectins bind only to the glycans of captured proteins. After this chemically blocking, serum samples were incubated with the antibody microarray, followed by the glycans detection by using different biotinylated lectins and GBPs, and visualized with Cy3-streptavidin. The parallel use of an antibody panel and multiple lectin probing provides discrete glycosylation profiles of multiple proteins in a given sample 18-20. This method has been used successfully in multiple different labs 1, 7, 13, 19-31. However, stability of MPBH and Cys-Gly, complicated and extended procedure in this method affect the reproducibility, effectiveness and efficiency of the method. In this new protocol, we replaced both MPBH and Cys-Gly with one much more stable reagent glutamic acid hydrazide (Glu-hydrazide), which significantly improved the reproducibility of the method, simplified and shorten the whole procedure so that the it can be completed within one working day. In this new protocol, we describe the detailed procedure of the protocol which can be readily adopted by normal labs for routine protein glycosylation study and techniques which are necessary to obtain reproducible and repeatable results.

In this study, we describe an improved protocol for a multiplexed high-throughput antibody microarray with lectin detection method that can be used in glycosylation profiling of specific proteins. This protocol features new reliable reagents and significantly reduces the time, cost, and lab equipment requirements as compared to the previous procedure.

 In this study, we describe an effective protocol for use in a multiplexed high-throughput antibody microarray with glycan binding protein detection that ...

Residue-Specific Exchange of Proline by Proline Analogs in Fluorescent Proteins: How "Molecular Surgery" of the Backbone Affects Folding and Stability

Replacement of proline (Pro) residues in proteins by the traditional site-directed mutagenesis by any of the remaining 19 canonical amino acids is often detrimental to protein folding and, in particular, chromophore maturation in green fluorescent proteins and related variants. A reasonable alternative is to manipulate the translation of the protein so that all Pro residues are replaced residue-specifically by analogs, a method known as selective pressure incorporation (SPI). The built-in chemical modifications can be used as a kind of &#34;molecular surgery&#34; to finely dissect measurable changes or even rationally manipulate different protein properties. Here, the study demonstrates the usefulness of the SPI method to study the role of prolines in the organization of the typical &#946;-barrel structure of spectral variants of the green fluorescent protein (GFP) with 10-15 prolines in their sequence: enhanced green fluorescent protein (EGFP), NowGFP, and KillerOrange. Pro residues are present in connecting sections between individual &#946;-strands and constitute the closing lids of the barrel scaffold, thus being responsible for insulation of the chromophore from water, i.e., fluorescence properties. Selective pressure incorporation experiments with (4R)-fluoroproline (R-Flp), (4S)-fluoroproline (S-Flp), 4,4-difluoroproline (Dfp), and 3,4-dehydroproline (Dhp) were performed using a proline-auxotrophic E. coli strain as expression host. We found that fluorescent proteins with S-Flp and Dhp are active (i.e., fluorescent), while the other two analogs (Dfp and R-Flp) produced dysfunctional, misfolded proteins. Inspection of UV-Vis absorption and fluorescence emission profiles showed few characteristic alterations in the proteins containing Pro analogs. Examination of the folding kinetic profiles in EGFP variants showed an accelerated refolding process in the presence of S-Flp, while the process was similar to wild-type in the protein containing Dhp. This study showcases the capacity of the SPI method to produce subtle modifications of protein residues at an atomic level (&#34;molecular surgery&#34;), which can be adopted for the study of other proteins of interest. It illustrates the outcomes of proline replacements with close chemical analogs on the folding and spectroscopic properties in the class of &#946;-barrel fluorescent proteins.

To overcome the limitations of classical site-directed mutagenesis, proline analogs with specific modifications were incorporated into several fluorescent proteins. We show how the replacement of hydrogen by fluorine or of the single by double bonds in proline residues ("molecular surgery") affects fundamental protein properties, including their folding and interaction with light.

Replacement of proline (Pro) residues in proteins by the traditional site-directed mutagenesis by any of the remaining 19 canonical amino acids is often ...

Rapid Antibody Glycoengineering in Chinese Hamster Ovary Cells

Recombinant monoclonal antibodies bind specific molecular targets and, subsequently, induce an immune response or inhibit the binding of other ligands. However, monoclonal antibody functionality and half-life may be reduced by the type and distribution of host-specific glycosylation. Attempts to produce superior antibodies have inspired the development of genetically modified producer cells that synthesize glyco-optimized antibodies. Glycoengineering typically requires the generation of a stable knockout or knockin cell line using methods such as clustered regularly interspaced short palindromic repeats (CRISPR)-associated protein 9. Monoclonal antibodies produced by engineered cells are then characterized using mass spectrometric methods to determine if the desired glycoprofile has been obtained. This strategy is time-consuming, technically challenging, and requires specialists. Therefore, an alternative strategy that utilizes streamlined protocols for genetic glycoengineering and glycan detection may assist endeavors toward optimal antibodies. In this proof-of-concept study, an IgG-producing Chinese hamster ovary cell served as an ideal host to optimize glycoengineering. Short interfering RNA targeting the Fut8 gene was delivered to Chinese hamster ovary cells, and the resulting changes in FUT8 protein expression were quantified. The results indicate that knockdown by this method was efficient, leading to a ~60% reduction in FUT8. Complementary analysis of the antibody glycoprofile was performed using a rapid yet highly sensitive technique: capillary gel electrophoresis and laser-induced fluorescence detection. All knockdown experiments showed an increase in afucosylated glycans; however, the greatest shift achieved in this study was ~20%. This protocol simplifies glycoengineering efforts by harnessing in silico design tools, commercially synthesized gene targeting reagents, and rapid quantification assays that do not require extensive prior experience. As such, the time efficiencies offered by this protocol may assist investigations into new gene targets.

The glycosylation pattern of an antibody determines its clinical performance, thus industrial and academic efforts to control glycosylation persist. Since typical glycoengineering campaigns are time- and labor-intensive, the generation of a rapid protocol to characterize the impact of glycosylation genes using transient silencing would prove useful.

Recombinant monoclonal antibodies bind specific molecular targets and, subsequently, induce an immune response or inhibit the binding of other ligands. ...

Targeting Cysteine Thiols for in Vitro Site-specific Glycosylation of Recombinant Proteins

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